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J Neurochem. Author manuscript; available in PMC 2016 Jan 1.
Published in final edited form as:
J Neurochem. 2015 Jan; 132(2): 169–182.
Published online 2014 Oct 31. doi: 10.1111/jnc.12967
PMCID: PMC4302046
NIHMSID: NIHMS638352
PMID: 25319522

P301L Tau Expression Affects Glutamate Release and Clearance in the Hippocampal Trisynaptic Pathway

Holly C. Hunsberger,1 Carolyn C. Rudy,1 Seth R. Batten,2 Greg A. Gerhardt,2 and Miranda N. Reed1,3,4,*
Author information Copyright and License information PMC Disclaimer

Abstract

Individuals at risk of developing Alzheimer’s disease (AD) often exhibit hippocampal hyperexcitability. A growing body of evidence suggests perturbations in the glutamatergic tripartite synapse may underlie this hyperexcitability. Here, we used a tau mouse model of AD (rTg(TauP301L)4510) to examine the effects of tau pathology on hippocampal glutamate regulation. We found a 40% increase in hippocampal vGLUT, which packages glutamate into vesicles, and has previously been shown to influence glutamate release, and a 40% decrease in hippocampal GLT-1, the major glutamate transporter responsible for removing glutamate from the extracellular space. To determine whether these alterations affected glutamate regulation in vivo, we measured tonic glutamate levels, potassium-evoked glutamate release, and glutamate uptake/clearance in the dentate gyrus (DG), CA3, and CA1 regions of the hippocampus. P301L tau expression resulted in a 4- and 7-fold increase in potassium-evoked glutamate release in the DG and CA3, respectively, and significantly decreased glutamate clearance in all 3 regions. Both release and clearance correlated with memory performance in the hippocampal-dependent Barnes maze task. Alterations in mice expressing P301L were observed at a time when tau pathology was subtle and before readily detectable neuron loss. These data suggest novel mechanisms by which tau may mediate hyperexcitability.

Keywords: Alzheimer, tau, glutamate clearance, in vivo electrochemistry, synaptic release, hippocampus

Introduction

Alzheimer’s disease (AD) affects approximately 5.4 million Americans and is often characterized by progressive memory loss, decline in cognitive skills, and adverse behavioral changes (Thies & Bleiler 2013). Biologically, Alzheimer’s disease is characterized by an abundance of extracellular amyloid plaques, comprised of aggregates of beta-amyloid (Aβ), and intracellular neurofibrillary tangles containing hyperphosphorylated tau protein (Serrano-Pozo et al. 2011). Another major feature of AD is neurodegeneration that may relate to neural network dysfunction (Seeley et al. 2009, Buckner et al. 2005, Palop et al. 2006). Recent work suggests tau-related pathology begins in vulnerable regions of the brain, including the entorhinal cortex and hippocampus, part of the brain’s memory network, before spreading to other cells along the same neural network (Liu et al. 2012, de Calignon et al. 2012, Nath et al. 2012).

Effective regulation of activity in these neural networks is essential because both increases and decreases in stimulation can impair neuronal function and survival, while neural network dysfunction could contribute directly to the neurodegenerative process (Palop et al. 2006). An early feature in aging, before AD pathology, is the hyperactivity of the memory network, particularly hippocampal regions. In studies using functional magnetic resonance imaging (fMRI), elevated hippocampal activation is observed in individuals at risk for AD, including cognitively normal carriers of the ApoE4 allele, a known genetic risk factor for AD (Filippini et al. 2009, Dennis et al. 2010, Trivedi et al. 2008), pre-symptomatic carriers of genetic mutations in familial AD (Quiroz et al. 2010), and patients with mild cognitive impairment (MCI) (Dickerson et al. 2005, Hamalainen et al. 2007). Longitudinal fMRI assessments of ApoE4 allele carriers indicate that hippocampal overactivation correlates with declines in memory (Bookheimer et al. 2000). Furthermore, patients with MCI exhibit greater hippocampal activation during memory encoding (Celone et al. 2006, Dickerson et al. 2004, Dickerson et al. 2005, Sperling 2007), and increased activation in MCI is predictive of the degree and rate of cognitive decline and for conversion to AD (Miller et al. 2008).

Hippocampal hyperactivity was once believed to serve as a compensatory function for deteriorating circuitry by recruiting extra neural resources (i.e., greater cognitive effort to achieve comparable performance) (Ward & Frackowiak 2003, Grady et al. 2003, Bondi et al. 2005). However, more recent studies show excess activation may contribute directly to memory impairment and AD-related pathology and could represent a therapeutic target. Circumstantial human evidence supports this view. For example, seizures and epileptiform activity are associated with an early age at onset of cognitive decline and precede or coincide with diagnosis of MCI or AD (Vossel et al. 2013). However, the relation between hyperactivity and memory impairments may be more than correlational (Koh et al. 2010, Sanchez et al. 2002, Bakker et al. 2012). Treatments targeting excess hippocampal activation dose-dependently improved memory performance in memory-impaired aged rats; these same doses had no effect in young rats without memory impairments, suggesting dampening of hippocampal hyperactivity, not merely a global cognitive enhancement, was responsible for the memory improvement in aged rats (Koh et al. 2010). Furthermore, a reduction in aberrant neural network activity reversed the synaptic and cognitive deficits observed in a mouse model of AD (Sanchez et al. 2012). Evidence for the adverse consequences of hyperexcitability has also been shown in humans; reducing hippocampal activation in an amnestic MCI group improved memory performance (Bakker et al. 2012). Together, these studies suggest increased hippocampal activation is not merely a compensatory response but a dysfunctional condition, and a condition that may be permissive for the development of AD.

Recent work suggests tau may mediate hyperexcitability. Genetic removal of tau decreases seizure activity in an Aβ mouse model of AD (Roberson et al. 2011). Furthermore, in this same mouse model, reducing endogenous tau ameliorated excitotoxicity and rescued cognitive dysfunction, without altering Aβ levels (Roberson et al. 2007), suggesting tau, not Aβ, was mediating excitotoxicity. Deletion of tau in mouse and drosophila models of epilepsy also reduces hyperexcitability, as well as seizure frequency and duration (Daniels et al. 2011). The exact mechanism for these changes remains to be determined, but recent work suggests tau may alter glutamate neurotransmission (Roberson et al. 2011, Roberson et al. 2007, Timmer et al. 2014).

To examine the role of tau in glutamate dysregulation, we used the most commonly used tau mouse model of AD, the rTg(TauP301L)4510 (hereafter called TauP301L) mouse model. These mice exhibit age-dependent cognitive decline, neurofibrillary tangle deposition, and neuron loss (Ramsden et al. 2005, SantaCruz et al. 2005). However, previous work suggests TauP301L mice exhibit electrophysiological hyperexcitability prior to tangle deposition or neuronal death (Crimins et al. 2012). Here, we sought to examine the effects of P301L expression on glutamate regulation at an age when subtle memory deficits and tau pathology are detectable but before neuron loss or tangle deposition occurs. This allowed us to dissociate the memory loss and any glutamate alterations resulting from P301L human tau expression with that associated with neuronal loss and to potentially provide an explanation for the electrophysiological hyperexcitability observed in this model.

Prior to examination of glutamate regulation, mice were memory tested using the hippocampal-dependent Barnes maze (BM) task to ensure the presence of subtle memory impairments at the age tested. In vivo glutamate regulation was measured in the DG, CA3, and CA1 subregions of the hippocampus, areas rich in glutamate receptors (Nimchinsky et al. 2004, Pettit & Augustine 2000), using ceramic-based microelectrode arrays (MEAs) coupled with amperometry. This approach allowed us to selectively measure extracellular glutamate at 10 Hz and compare tonic glutamate levels, potassium-evoked (KCl) glutamate release, and glutamate uptake/clearance among the groups (Burmeister & Gerhardt 2001, Burmeister et al. 2000). Another benefit of MEAs over other ex vivo methods is the ability to study brain regions in vivo without disrupting their extrinsic and intrinsic connections, a particularly important consideration when examining the complex connections of the trisynaptic loop of the hippocampus. Because examination of memory and glutamate regulation was done within the same animal, we were able to relate changes in glutamatergic signaling with behavioral changes. The current study suggests targeting excess hippocampal activity may have therapeutic potential for the treatment of AD.

Methods

Subjects

Creation of TauP301L mice has been described (Hoover et al. 2010, Ramsden et al. 2005, SantaCruz et al. 2005). Briefly, regulatable transgenic mice expressing human four-repeat tau lacking the N-terminal sequences (4R0N) with a P301L mutation were created by crossing a responder and activator line. Responder mice (FVB/N background strain), heterozygous for the TRE-TauP301L transgene, were bred with heterozygous activator mice (129S6 background strain) that express the tet-off tetracycline transactivator (tTA) reading frame placed downstream of Ca2+/calmodulin kinase II (CaMKII) promoter elements (SantaCruz et al. 2005). The four genotypes of animals generated are described by the following nomenclature, TauP301L/CKtTA, and include what we refer to as TauP301L mice (+/+) and Controls (−/+, +/−, −/−). In the current study, we used the −/+ control because no behavioral differences among the 3 Control groups have been reported, and human tau is not expressed in any of the Controls (Ramsden et al. 2005, SantaCruz et al. 2005, Hoover et al. 2010), yet the −/+ allowed us to control for the expression of tTA (Mayford et al. 1996).

To control for the overexpression of human tau, we also examined the rTg(TauWT) 21221 mouse model (hereafter called TauWT). TauWT mice express wild-type 4R0N human tau at concentrations equivalent to P301L human tau in TauP301L mice, but without the P301L mutation, and show no neurofibrillary tangles, progressive memory decline nor neurodegeneration (Hoover et al. 2010). TauWT were generated in the same way as TauP301L mice.

The CaMKII promoter was used to restrict TauP301L protein expression to the forebrain (SantaCruz et al. 2005), which contains the brain regions most severely affected in AD (Auld et al. 2002), while the tet-off system allowed for regulatable expression of the tau protein. In the tet-off system, the tTA protein binds to TRE in the absence of doxycycline, allowing transcription and protein expression to occur (Liu et al. 2008), whereas in the presence of doxycycline, the tTA protein cannot bind to TRE, allowing tau expression to be suppressed. Previously published work suggests developmental P301L tau expression produces alterations not observed following adult-onset tau expression (Caouette et al. 2013), possibly due to the important role of tau in brain development (Wang & Liu 2008). To avoid mutant tau expression during the perinatal and early postnatal stages, thereby preventing developmental alterations unrelated to AD, tau was suppressed during brain development (Hunsberger et al. 2014). To suppress tau, 40 ppm doxycycline hyclate (DOX) was administered via water bottles to breeder dams for three weeks prior to mating and to all experimental female mice from birth until 2 months of age (Hölscher 1999). Behavioral testing began at 5 months of age, after 3 months of tau expression, and was followed by glutamate function testing. All experimental procedures were conducted in accordance with the standards of International Animal Care and Use Committee, and the West Virginia University Animal Care and Use Committee approved all experimental procedures.

Barnes Maze

The Barnes maze consists of a circular white platform (122 cm), 108 cm above the ground, with 40 holes (5 cm), one with a hidden escape box. Training was performed as previously described with slight modification (McAfoose et al. 2009, McLay et al. 1998). For the two days preceding acquisition trials, the mice were habituated to the maze. There were no extra-maze cues during habituation training. During the first day of habituation, the mice were placed under a clear beaker and allowed to enter the escape box for 5 min. The next day, the mouse was gently guided to the escape box and allowed to remain in the escape box for 2 min. Each mouse completed two trials each day with approximately 20–25 min between trials.

For acquisition training on days 3–7, extra-maze cues were placed around the room, and weak aversive stimulation was applied to increase the motivation to escape from the circular platform; aversive stimulation included the illumination of overhead lights and the use of four fans evenly placed around the maze. At the start of the trial, the mouse was placed under a plastic beaker in the center of the platform. After 10 sec, the beaker was raised, and the mouse was free to explore. The trials ended 30 sec after the mouse entered the escape box. If the mouse did not enter the escape box during the 3-min trial, the experimenter gently guided the mouse into the escape box and covered the hole for 30 sec. Mice received 3 trials per day with 20–25 min between trials. The maze was cleaned with 70% ethanol, rotated clockwise after every trial to avoid intra-maze odor or visual cues, though the escape box remained in the same place relative to extra-maze cues. Latency to reach the escape box and number of errors before reaching the escape box were recorded.

The first of two probe trials, in which the escape box was removed, took place 24 hr after the last acquisition trial. The second probe trial took place one week following the first probe trial. Probe trials lasted 90 sec. Latency to reach the target hole and the number of errors made were recorded. Because of the long duration of the probe trial and to further evaluate search strategy, as previously described (e.g., Devan et al. 2003), the probe trial was divided into three 30 sec epochs, and errors evaluated in each epoch separately.

Enzyme-based microelectrode arrays

Ceramic-based MEAs were used to examine glutamate regulation and were purchased from Quanteon, L.L.C. (Nicholasville, KY). The array consisted of a ceramic-based multisite microelectrode with 4 platinum recording sites (Burmeister & Gerhardt 2001). These sites were arranged in dual pairs to allow interfering agents to be detected and removed from the analyte signal (Burmeister & Gerhardt 2001). Coating of the microelectrodes has been described previously (Hinzman et al. 2010). Briefly, the recording sites were covered with glutamate-oxidase (GluOx) to oxidize glutamate to alpha-ketoglutarate and hydrogen peroxide (H202), the reporter molecule (Burmeister & Gerhardt 2001). An inactive protein matrix covered the other pair of recording sites (sentinel sites). Small molecules, like H202, can diffuse through the mPD exclusion layer, but not larger molecules such as ascorbic acid or monoamines. The background current from the sentinel sites was then subtracted from the recording sites to produce a selective measure of extracellular glutamate. A reference electrode Ag/AgCl was implanted into a remote site from the recording area (Burmeister & Gerhardt 2001).

Calibration

Calibrations were conducted on the MEAs prior to their use to ensure sensitivity and selectivity and to create a standard curve for the conversion of current to glutamate concentration. Using the FAST-16 mkII system (Quanteon), a constant potential of + 0.7 V versus an Ag/AgCl reference was applied to the MEA to oxidize the reporter molecule. The resulting current was amplified, digitized, and filtered by the FAST 16 mkII system. The MEA tip was submerged in 40 mL of a 0.05 M phosphate-buffered saline (PBS) maintained at 37°C. A standard curve was determined by adding successive aliquots of 20 µL glutamate to achieve concentrations of 20, 40, and 60 µM. The increase in current (nA) produced by oxidation was used to calculate the calibration slope to a known concentration of glutamate (Burmeister, et al., 2002). To determine selectivity for glutamate, ascorbic acid (250 µM) and dopamine (2 µM) were added to the solution (Hinzman et al. 2012). To determine the limit of detection (LOD), the smallest concentration of glutamate that can be measured by the device, the slope of the standard curve was used, as well as the noise or relative standard deviation of the baseline signal (Hinzman et al. 2012).

MEA/Micropipette Assembly

For intracranial drug deliveries, a glass micropipette with an inner diameter tip of 10–15 µm (Quanteon) was attached to the MEA. The micropipette was centered between the dorsal and ventral platinum recording pairs and positioned 80–100 µm away from the MEA surface. Location of the micropipette to the MEA was verified post-surgery to ensure that the pipette did not move. The micropipette was back-filled with sterile-filtered isotonic KCl solution (70 mM KCl, 79 mM NaCl, 2.5 mM CaCl2, pH 7.4) or glutamate solution (200 µM glutamate, pH 7.4). The micropipette was attached to a Picospritzer III (Parker-Hannifin, Cleveland, OH) and set to consistently deliver volumes of 50–100 nL. Pressure was applied from 2 – 20 psi (0.138 – 1.38 bar; 13.8 – 137.8 kPa) for .30 – 2.5 sec. Volume displacement was monitored with the use of a stereomicroscope fitted with a reticule (Friedemann & Gerhardt 1992).

In vivo Anesthetized Recordings

Mice were anesthetized with isoflurane (1–4% inhalation; continuous) and placed into a stereotaxic apparatus (David Kopf Instruments, Tujunga, CA, USA). Isoflurane was used because other anesthetics have been shown to have great effects on resting glutamate levels (Mattinson et al. 2011). Though initial reports suggested isoflurane increases tau phosphorylation (Planel et al. 2004), more recent reports suggest that when anesthesia-induced hypothermia is controlled for, isoflurane does not increase tau phosphorylation (Tan et al. 2010). To ensure our mice did not become hypothermic while under anesthesia, body temperature was continuously measured using a rectal probe and maintained at 37°C with a water pad connected to a recirculating water bath (Gaymar Industries Inc., Orchard Park, NY).

A craniotomy was performed to allow access to the hippocampus for MEA recordings. The Ag/AgCl wire reference electrode was placed under the skin with a saline-soaked gauze pad in the hemisphere opposite from the recording sites. The MEA/micropipette array was placed into the DG, CA3, and CA1 of the hippocampus, sub-regions rich in glutamate receptors (Nimchinsky et al. 2004, Pettit & Augustine 2000). Stereotaxic coordinates for the different sub-regions of the hippocampus were calculated using the mouse brain atlas (Paxinos & Watson 2004) [DG (AP: −2.3 mm, ML: +/−1.5 mm, DV: 2.1 mm), CA3 (AP: −2.3 mm, ML: +/−2.7 mm, DV: 2.25 mm), CA1 (AP: −2.3 mm, ML: +/−1.7 mm, DV: 1.4 mm)] and confirmed post mortem. All MEA recordings were performed at 10 Hz using constant potential amperometry recordings with the FAST-16. After the MEA reached a stable baseline (10–20 min), tonic glutamate levels (µM) were calculated averaging extracellular glutamate levels over 10 seconds prior to any application of solutions. In all three subregions of one hemisphere, evoked release (i.e., amplitude) was measured by local application of KCl delivered every 2–3 minutes. KCl-evoked release of glutamate is highly reproducible and indicative of the intact glutamate neuronal system that is detected by the MEAs (Day et al. 2006). After 10 reproducible signals, the results were averaged for each group and the average amplitude compared (Hinzman et al. 2012, Nickell et al. 2007, Hinzman et al. 2010). KCl-evoked release of glutamate was measured to determine the “capacity” of the nerve terminals to release glutamate (Hinzman et al. 2010).

To examine glutamate clearance/uptake, exogenous glutamate was applied in the opposite hemisphere. After the MEA reached a stable baseline (10–20 min), varying volumes of 200 µM sterile-filtered glutamate solution were applied into the extracellular space every 2–3 minutes. Glutamate signals with amplitudes of 40 µM or below were analyzed as this was the physiological range of KCl-evoked glutamate release observed in the current study and previous studies (Hinzman et al. 2010). The net area under the curve (AUC) was used to estimate glutamate clearance. The hemispheres used for KCl and glutamate application were counterbalanced, as was the order of sub-regions within a hemisphere. After recording from all locations, an MEA with an attached micropipette was used to locally apply Fluoro-Ruby (Millipore) or fast green (Sigma), which was used to confirm MEA placement following brain sectioning (Figure 1). Prior studies have shown that the MEAs produce minimal effects both acutely and chronically (Hascup et al. 2009). All behaviorally tested mice underwent glutamate examination. However, data from some hippocampal regions were excluded for reasons including death during surgery, poor placement of the MEA, or in the case of the glutamate uptake studies, amplitudes greater than 40 uM. For each measure, the number of mice per group is provided in the corresponding figure caption.

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Cresyl violet-stained 20 µm section of hippocampus shows location of MEA tracks in CA3 and CA1.

Immunoblotting

To ensure application of KCl or exogenous glutamate did not influence protein expression, hippocampal tissues from mice not used for glutamate testing were used to assess vGLUT1 and GLT-1 expression. Hippocampal tissue was prepared for immunoblotting using 500 ul RIPA extraction buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 0.5% Triton X-100, 1 mM EDTA, 3% SDS, 1% Na deoxycholate) with Roche protease inhibitor tablets (EDTA-free Easy pack tablets, Phosphostop Easy pack tablets) added fresh the day of extraction. Protein concentrations were determined with a BCA protein assay using BSA as a standard.

Hippocampal tissue samples were thawed and 10 µg aliquots were mixed with loading buffer (450 mM Tris HCL, pH 8, 8% SDS, 24% glycerol, 5% mercapoethanol, 0.1% bromophenol blue, 0.1% phenol red). Before loading, samples were either heated to 70°C (vGLUT1 and synaptophysin) or 95°C (GLT-1 and actin I-19) for 5 min and then separated on 10% criterion gels (Biorad, #345-0009), and transferred onto .45 µm polyvinylidene difluoride membranes (Millipore, Bedford, MA). Membrane blots were blocked for 1 h at RT in 5% BSA in 0.1% Tween 20/Tris-buffered saline (TTBS; 198.5 mM NaCl, 9.98 mM Trizma base, pH 7.4) (vGLUT-1, synaptophysin, and actin) or 5% milk in TTBS (Non-fat dry milk, Cell Signaling) (GLT-1). After blocking, membranes were incubated with an antibody directed against the protein of interest (vGLUT1, 1:6,000; GLT-1, 1:10,000, synaptophysin, 1:200; actin I-19, 1:500) overnight at 4°C. The next day, membranes were incubated with Streptactin-HRP (Biorad) and the appropriate biotinylated or HRP-conjugated secondary antibody for 1.5 h at RT. Blots were then sprayed 5–10 times evenly with Rapidstep ECL (Calbiochem), incubated for 5 minutes, and visualized using Fluorchem E imager (Cell Biosciences). Membranes used to detect vGLUT1 or GLT-1 signals were first probed with vGLUT1 or GLT-1 antibody, then stripped for 1 h with Restore PLUS western blot stripping buffer (Peirce Chemical) and re-probed with synaptophysin or actin antibody, respectively. vGLUT and GLT-1 bands were normalized to synaptophysin or actin, respectively. Band density was measured using AlphaView software (Proteinsimple, Santa Clara, California, USA).

Immunohistochemistry

Immunohistochemical detection of total and phosphorylated tau species in transgenic and control mice was performed as previously described (Hoover et al. 2010, Ramsden et al. 2005). To ensure application of KCl or exogenous glutamate did not influence tau phosphorylation, tissue from mice not used for glutamate testing was utilized for IHC. Briefly, hemibrains were immersion fixed in 10% formalin for 24–48 h and embedded in paraffin. Serial sections were cut at 5 µm using a microtome, mounted onto CapGap slides (Thermo-Fisher), and rehydrated according to standard protocols. Mounted slides were pretreated with a citrate buffer (pH 6.0) in a Black & Decker (Owings, MD) steamer for 30 min, with a 10 min cool down. Standard 2 d immunostaining procedures using peroxidase-labeled streptavidin and DAB chromagen on an automated TechMate 500 capillary gap immunostainer (Ventana Medical Systems, Tucson, AZ) were used with antibodies directed against (Table 1). Hematoxylin counterstaining was used to provide cytological detail. Hematoxylin and eosin and modified Bielschowsky silver staining to detect neurofibrillary tangles (NFTs) were performed using standard histological techniques. Photomicrographs of hippocampal and cortical neurons were captured at three different magnifications (×5, ×10 and ×40) with a Zeiss Axioskop microscope coupled to a CCD camera and processed and assembled in Adobe Photoshop. No positive labeling was observed for pathological tau epitopes in nontransgenic mice.

Table 1

Tau Antibodies.

AntibodySpeciesSpecificityUseDilutionSource
MC-1MouseTau; conformational epitope, 7–9 and 326–330 aaConformation specific; early pathological change in AD1:8000P.Davies
CP-13MouseTau; pSer202Phosphorylation specific; early pathological change in AD1:2000P.Davies
AT-8MouseTau; pSer202/pThr205Pre-tangles; mid-stage1:4000Innogenetics
PG-5MouseTau; pSer409Pre-tangles; mid-stage1:200P.Davies
NeuNMouseNeuNNeuronal marker1:4000Chemicon

Data analysis

All statistical analyses were performed using JMP (SAS, Cary, NC 27513). Statistical analysis consisted of ANOVA and repeated-measures ANOVA (RMANOVA), followed by Tukey post hoc comparisons. For the RMANOVA of behavioral data, transgene status served as the between-subject variable, and session served as the within-subject variable. Amperometric data were analyzed using a custom Microsoft excel software program (MatLab). To determine concentrations of glutamate in the hippocampus, the background current from the sentinel sites was subtracted from the signal obtained from the GluOx recording sites. The resting current (pA) was divided by the slope (µM/pA) obtained during calibration and reported as a concentration of glutamate. Using Pearson r correlations, KCl-evoked glutamate release (amplitude) and glutamate clearance (net AUC) in the DG, CA3, and CA1 were correlated separately with performance in the Barnes maze (errors). Correlations were run only for those mice in which data for both behavior and glutamate were analyzed.

Results

Alterations in hippocampal vGLUT1 and GLT-1 expression in TauP301L mice

Hippocampal vGLUT1 expression was significantly increased in TauP301L mice [F(2,14) = 8.65, p = .004]. To determine if there was a widespread increase in presynaptic terminals and to serve as a loading control, synaptophysin immunoblotting was performed. No statistically significant difference in synaptophysin expression was observed among the groups [F(2,14) = 1.94, p = .18]. The ratio of vGLUT1 to synaptophysin was also significantly increased in TauP301L mice [F(2,14) = 8.08, p =.005] (Figure 2A).

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TauP301L mice exhibit increased vGLUT1 expression and decreased GLT-1 expression in the hippocampus

(A) vGLUT1 expression was significantly increased in hippocampal tissue of TauP301L mice. (B) In contrast, GLT-1 expression was significantly decreased in TauP301L mice. (Mean ± SEM; *p<.05 Control vs. TauP301L, # p< .05 TauWT vs. TauP301L, ## p< .01 TauWT vs. TauP301L, n = 5–6/group).

Comparison of hippocampal GLT-1 expression revealed a significant decrease in the hippocampus of TauP301L mice [F(2,13) = 4.94, p = .03], but no differences among the groups for the loading control, β-actin [F(2,13) = 0.56, p = .59]. The ratio of GLT-1 to β-actin was also significantly lower in the hippocampus of TauP301L mice [F(2,13) = 6.59, p = .01] (Figure 2B).

Spatial learning and memory deficits in TauP301L mice

During acquisition, latency did not differ among the groups [Tg main effect: F(2,25) = 1.45, p = .25; Tg*Day interaction: F(8,100) = 1.39, p = .21]. However, TauP301L mice made significantly more errors during acquisition [Tg main effect: F(2,25) = 4.32, p = .03], regardless of day [Tg*Day interaction: F(8,100) = 1.64, p = .12] (Figure 3A). Examination of the 24 hour probe trial in 30 sec epochs, as previously reported (e.g., Devan et al. 2003), indicated TauP301L mice made significantly more errors in the 30–60 sec epoch [F(2,23) = 3.98, p = .03)] (Figure 3B). When the total errors in the 24 hour probe were compared, TauP301L mice made marginally more errors across the entire 90 sec probe trial [F(2,23) = 3.13, p = .06)] (Figure 3C). For the 1 week probe trial, there were no differences among the groups for total errors, errors in the three epochs, or latency (ps > .05; data not shown).

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TauP301L mice exhibit memory deficits in the Barnes maze task

(A) Errors during acquisition were significantly increased for TauP301L mice. (B) TauP301L mice made more errors during the 30–60 sec epoch. (C) TauP301L mice exhibited marginally more errors during probe trials. (Mean ± SEM; *p<.05 Control vs. TauP301L, ** p< .01 Control vs. TauP301L, # p< .05 TauWT vs. TauP301L, n = 7–11/group; Controls = 7, TauWT = 11, and TauP301L = 9).

TauP301L mice exhibit hippocampal glutamate dysregulation

Tonic glutamate levels were not significantly different in the DG [F(2,17) = .15; p = .86], CA3 [F(2,16) = .11; p = .89], or CA1 [F(2,17) = .58; p = .57] among the Controls, TauWT, or TauP301L mice (Figure 4A). To examine the capacity for glutamate release, KCl was delivered via a micropipette. Local application of 50–100 nL of 70 mM KCl produced reproducible glutamate release in all regions of the hippocampus. The amplitudes of KCL-evoked-glutamate release in the CA1 [F(2,18) = 1.31; p = .29] were similar among the groups, whereas in the DG [F(2,17) = 4.14; p = .03] and CA3 [F(2,17) = 4.33, p = .03], the amplitude of KCl-evoked glutamate release was 4 and 7 times larger in TauP301L mice, respectively (Figure 4B,C).

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Tonic and potassium chloride (KCl)-evoked release of glutamate in the DG, CA3, and CA1 regions of the hippocampus

(A) Tonic glutamate levels were not significantly different in the hippocampus. (B) Baseline-ded representative recordings of KCl-evoked glutamate release in the CA3 showed that there was a significant increase in the amplitude of glutamate release in TauP301L mice. Local application of KCl (↑) produced a robust increase in extracellular glutamate that rapidly returned to tonic levels. (C) The average amplitudes of KCl-evoked glutamate release in the DG and CA3 regions of hippocampus were significantly increased in the TauP301L mice after local application of 50–100 nL of 70 mM KCl. (Mean ± SEM; *p<.05 Control vs. TauP301L; # p< .05 TauWT vs. TauP301L; n = 6–7/group).

Rapid application of glutamate into the extracellular space allowed us to mimic endogenous glutamate release and examine glutamate clearance back to baseline in vivo. To ensure differences in net area under the curve (AUC) among the groups following application of endogenous glutamate were due to alterations in clearance and not differences in amount of endogenous glutamate applied, we first compared the amplitude of glutamate signals following administration of exogenous glutamate; no differences in amplitude were observed among Controls, TauWT, and TauP301L mice in the DG [F(2,14) = .88; p = .44], CA3 [F(2,12) = .47; p = .64], and CA1 [F(2,14) = .04; p = .96] (Figure 5A), suggesting similar application of exogenous glutamate. We next examined Trise, the time for signal to reach maximum amplitude, to determine if transgene status altered diffusion of glutamate in the extracellular space; Trise was not significantly different among the groups DG [F(2,14) = .33; p = .72], CA3 [F(2,12) = .19; p = .83], and CA1 [F(2,14) =.58; p = .57] (Figure 5B), suggesting any reductions in glutamate clearance were not due to diffusion from the point source (micropipette) to the MEA (Sykova et al. 1998). Because neither amplitude nor Trise differed among the groups, any differences in net AUC likely results from decreases in glutamate uptake. Following exogenous application of glutamate, TauP301L mice exhibited an increased net AUC in the DG [F(2,14) = 5.21, p = .02], CA3 [F(2,12) = 7.37; p = .008], and CA1 [F(2,14) = 8.75; p = .003] (Figure 5C,D), suggesting reduced glutamate uptake in all 3 regions of the hippocampus.

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Reduced glutamate clearance in TauP301L mice following exogenous glutamate application in the DG, CA3, and CA1 regions of the hippocampus

(A) The amplitude of glutamate signal was similar among groups in each region. (B) Trise, an indicator of glutamate diffusion, was similar among the groups in each region. (C) Representative glutamate signals in the CA1 from local application of 200 µM glutamate (↑) in Controls (circles), TauWT (squares), and TauP301L (triangles) mice. TauP301L mice showed significant decreases in net area under the curve (AUC). (D) The net AUC was increased in TauP301L mice in all 3 regions of the hippocampus in TauP301L mice, indicating reduced glutamate uptake. (Mean ± SEM; *p<.05 Control vs. TauP301L, # p< .05 TauWT vs. TauP301L; n = 6–7/group).

Glutamate regulation correlates with Barnes maze performance

We next sought to determine whether errors in the Barnes maze correlated with glutamate regulation (Table 2). For KCl-evoked release, the amplitude of evoked glutamate release in the CA3 was significantly correlated with Barnes maze performance (p = .002), whereas for the DG and CA1 regions, there was no relation between KCl-evoked release and performance (p = .21 and p = .21, respectively). In contrast, for clearance of exogenous glutamate, the opposite pattern was observed. Errors in the Barnes maze were significantly correlated with clearance (net AUC) in the DG (p = .0004) and CA1 (p = .002) but not the CA3 (p = .067).

Table 2

Correlations between Glutamate Dysregulation & Errors in the Barnes Maze

DGCA3CA1
KCl-evoked Release vs. ErrorsErrors=16.83+.58*Amp
r (21)= .29, p = .21		Errors=16.14+.024*Amp
r (20)= .64, p = .002		Errors=17.04+.38*Amp
r (20)= .29, p = .21
Clearance vs. ErrorsErrors=14.97+.064*AUC
r (19)= .76, p = .0004		Errors=15.67+.039*AUC
r (18)= .48, p = .067		Errors=15.07+.053*AUC
r (18)= .69, p = .002

Pathological tau conformation and phosphorylation

A panel of antibodies (Table 1) directed at biochemical changes in tau associated with AD was used to determine the extent of tau pathology at the time of behavioral and glutamate testing. As previously reported (Ramsden et al. 2005, Andorfer et al. 2003, Hunsberger et al. 2014), the earliest positive labeling in the hippocampus was identified using CP-13 and MC-1, which detect phosphorylation and conformation-specific changes, respectively (Figure 6 A,B). In contrast, when AT-8 and PG-5 were used to examine the presence of pretangles (accumulations of nonargyrophilic hyperphosphorylated tau in the neuronal cell body) in the hippocampus, no positive staining was observed with either antibody (Figure 6 C,D). Similarly, no NFTs were observed after staining with Bielschowsky silver (data now shown). To determine if there was readily detectable neuron loss, hematoxylin and eosin (H&E) stained tissue among the groups was compared. To further supplement the examination of neuronal loss, tissue was also incubated with a neuron-specific antibody (NeuN). After 3 months of tau expression, there was no readily detectable cell loss in the hippocampus of tauP301L mice using either H&E or NeuN (Figure 7).

An external file that holds a picture, illustration, etc. Object name is nihms638352f6.jpg
Early tau pathology in TauP301L mice

(A) IHC studies revealed abnormal tau conformation and phosphorylation when using the early pathological markers of AD, CP-13 and MC-1. (B) No differences were observed when the pretangle marker AT-8 and PG-5 were examined Conformational changes determined using: MC-1, amino acids 7 to 9 and amino acids 326 to 300, and phosphorylation changes detected using: CP-13, pSer202; AT-8, pSer202/pThr205; PG-5, pSer409. No positive labeling was observed after parallel processing of control tissue or TauWT mice (n = 4/group). Staining was consistent across all TauP301L mice (n = 4). Boxes indicate areas shown at higher magnification. Original magnifications: ×5 for hippocampus (Hippo.) and ×40 for CA1.

An external file that holds a picture, illustration, etc. Object name is nihms638352f7.jpg
No neuron loss was readily detectable in TauP301L mice

Neuron loss was not readily detectable when compared using (A) hematoxylin and eosin (H&E) or a (B) neuron-specific antibody (NeuN). Boxes indicate area shown at higher magnification. Representative images shown and were consistent across all TauP301L mice (n = 4). Original magnifications: ×5 for hippocampus (Hippo.) and ×40 for CA1.

Discussion

We examined the extent of glutamate dysregulation in a TauP301L mouse model of AD known to exhibit electrophysiological hyperexcitability prior to tangle deposition or neuronal death (Crimins et al. 2012). The present study is the first to use in vivo amperometry to examine glutamate dysregulation in the DG, CA3, and CA1 subregions of the hippocampus in an AD mouse model. The hippocampal-dependent Barnes maze task was used to relate pathological glutamate changes with functional deficits. Though tonic glutamate levels were unaltered in all three regions of the hippocampus, KCl-evoked glutamate release in the DG and CA3 was significantly increased in TauP301L mice, and glutamate clearance was significantly decreased in all 3 regions. This increase in release and decrease in clearance was associated with an increase in vGLUT1 and a decrease in GLT-1 expression, respectively.

The Barnes maze is a spatial memory task that requires animals to use spatial cues around the room to learn the position of a hole that can be used to escape the brightly lit, aversive, open surface of the maze. Rodents with hippocampal damage show impaired performance in the Barnes maze, suggesting this is a hippocampal-dependent task (Fox et al. 1998, Pompl et al. 1999). Unlike the commonly used hippocampal-dependent Morris water maze, the Barnes maze task does not require the mice to swim and is therefore considered less anxiogenic (Harrison et al. 2009), a notion supported by findings of lower plasma corticosterone in rodents examined after the Barnes maze compared to the Morris water maze task (Harrison et al. 2009). Because stress is related to both an increase in extracellular glutamate (Popoli et al., 2012) and an increase in tau hyperphosphorylation (Sotiropoulos et al. 2011, Popoli et al. 2012), the Barnes maze task was identified as a more suitable measurement of memory for the current study. The present findings confirm that the Barnes maze is a sensitive task for use with TauP301L mice, capable of detecting the subtle differences resulting from a short duration of P301L tau expression.

The tonic glutamate levels observed in the current study (1–3 µM) were similar to the levels previously reported in the hippocampus of microdialysis studies using rats (1–4 µM) (Lerma et al. 1986, Herman & Jahr 2007, Miele et al. 1996). Interestingly, in vitro studies using hippocampal brain slices suggest much lower ambient glutamate concentrations, with values closer to 0.025 µM (Herman & Jahr 2007). The reason for these discrepancies between in vitro and in vivo studies is unclear and warrants further investigation.

P301L tau expression did not alter tonic glutamate levels. The lack of differences in tonic glutamate for TauP301L mice is somewhat surprising given the increased vGLUT1 expression and KCl-evoked release observed in these mice. However, in vitro studies using hippocampal slices suggest tonic glutamate levels are not dependent on, or altered by, vesicular glutamate release but may instead be due to differences in glia-dependent release of glutamate (Jabaudon et al. 1999, Cavelier & Attwell 2005, Le Meur et al. 2007). In vivo studies using MEAs provide indirect support for this idea. In mice with traumatic brain injury, striatal tonic levels were increased, yet KCl-evoked released remained unchanged (Hinzman et al. 2010). MEA research with aging rats shows that the opposite is also possible; aged rats exhibited increased KCl-evoked release, but unaltered levels of tonic glutamate (Stephens et al. 2011), similar to the present findings. Future studies are needed to further delineate and confirm glia as primary sources of tonic extracellular glutamate.

The exact mechanism by which P301L tau expression increases KCl-evoked glutamate release in the DG and CA3 is not known, but the increased VGLUT expression observed in TauP301L mice might explain the increase in glutamate release. The number of VGLUT molecules has a direct impact on the number of glutamate molecules released by a single synaptic vesicle during exocytosis (Herzog et al. 2006, Wilson et al. 2005), and overexpression of VGLUT results in increased glutamate release, leading to excitotoxic neurodegeneration and a shortened lifespan (Daniels et al. 2011). An important future direction is to determine whether the increase in vGLUT1 expression in TauP301L mice is limited to the DG and CA3 regions of the hippocampus and whether reducing vGLUT1 expression attenuates the increased release observed in TauP301L mice.

In many neurodegenerative diseases, including amyotrophic lateral sclerosis (ALS) (Rothstein et al. 1995), Huntington’s disease (Arzberger et al. 1997), Parkinson’s disease (Ferrarese et al. 1999), and Alzheimer’s disease (Masliah et al. 2000), astrocytic glutamate transporter expression is decreased. However, because brains are often examined at the end-stage of these diseases, neuronal loss is often extensive, making it difficult to discern the role of glutamate transporters in the pathogenesis of these diseases. To better examine the role of glutamate transporters in the etiology of AD, mouse models have been examined. In JNPL-c tau transgenic mice, which also express the P301L mutation in tau, glutamate transporter (GLT-1) expression was increased by about 25% and was associated with improved stroke outcome (Liao et al. 2009). Surprisingly, we observed exactly the opposite effect of P301L tau expression: a decrease in GLT-1 expression. There are a few differences between the two studies that may account for this discrepancy. First, the previous finding of increased GLT-1 expression was observed after only 2 months of tau expression (Liao et al. 2009), whereas mice in the present study expressed tau for longer. Previous work with another P301L model suggests 2 months of P301L tau expression, before hyperphosphorylation such as that observed in the current study is present, results in increased long-term potentiation and improved cognitive performance (Boekhoorn et al. 2006). Because Liao and colleagues did not examine memory, it is unclear whether the increased GLT-1 expression was associated with improved cognitive performance. It is possible GLT-1 expression increases and then decreases with longer durations of tau expression. A second difference involves the age of onset of P301L tau expression. Mice in the previous study expressed tau during development (Liao et al. 2009), and thus, it is possible the increase in GLT-1 resulted from developmental tau expression. Previously published work suggests developmental P301L tau expression produces alterations not observed following adult-onset tau expression (Caouette et al. 2013). Such a finding is not surprising given the importance of tau in brain development (Wang & Liu 2008). Examination of GLT-1 at earlier and later time points in the adult-onset P301L model would help address these issues.

One potential caveat of the current study concerns the high spatial resolution of the microelectrodes (Burmeister et al. 2002, Burmeister et al. 2000). Because of the high spatial resolution, it is possible that different coordinates within the various subregions of the hippocampus would result in different effects than those observed here. In fact, when examined using MEAs, tonic glutamate levels were increased in rats with traumatic brain injury but only at certain depths within the striatum (Hinzman et al. 2010) increases in tonic glutamate levels were observed at a depth of 4.0 mm, but not at 4.5 mm or 5.0 mm. Thus, examination of TauP301L mice using slightly different coordinates might reveal different results. Though the spatial resolution can be a limitation, it is also a benefit, allowing subregional analyses and measurements of fast transmission close to the synapse.

In conclusion we used MEAs in combination with the TauP301L mouse model to examine the effects of P301L tau expression on glutamate signaling, without other mediators, such as Aβ plaques, tangles, and neuronal loss. Our results demonstrate memory-impaired TauP301L mice exhibit an increase in KCL-evoked glutamate release in the DG and CA3 regions of the hippocampus and a decrease in glutamate clearance in the DG, CA3, and CA1 regions of the hippocampus. More research is needed to determine the specific mechanisms by which tau pathology alters glutamate regulation.

Acknowledgements

This work was supported by the National Institute of General Medical Sciences (Reed - U54GM104942), the Alzheimer's Association (Reed - NIRG-12-242187), a WVU Faculty Research Senate Grant, and a WVU PSCOR grant. The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH or Alzheimer's Association. GG is the sole proprietor of Quanteon, LLC that makes the Fast-16 recording system used for glutamate measurements in this study.

Abbreviations

ADAlzheimer’s Disease
DGDentate gyrus
CA3Cornu Ammonis 3
CA1Cornu Ammonis 1
AβAmyloid- Beta
ECextracellular
MEAmicroelectrode array
LTPlong-term potentiation
tTATetracycline-Controlled Transcriptional Activation
TREtetracycline response element
BMBarnes Maze
mPDm-Phenylenediamine
GluOxglutamate oxidase
vGLUT-1vesicular glutamate transporter 1
GLT-1glutamate transporter 1
H&Ehematoxylin and eosin
NFTsneurofibrillary tangles

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